I. INTRODUCTION

Section:

ChooseTop of pageABSTRACTI. INTRODUCTION <<II. EXPERIMENTALIII. RESULTS AND DISCUSSI...IV. CONCLUSIONSUPPLEMENTARY MATERIALREFERENCESMicrofluidic technology provides the tools and techniques to handle and manipulate small volumes of fluids (Whitesides, 200649. Whitesides, G. M., “The origins and the future of microfluidics,” Nature 442(7101), 368–373 (2006). https://doi.org/10.1038/nature05058). Its advantages such as low reagent use, low response time, and high accuracy make microfluidic devices promising tools for the developing of point-of-care technologies (Pandey et al., 201840. Pandey, C. M., Augustine, S., Kumar, S., Kumar, S., Nara, S., Srivastava, S., and Malhotra, B. D., “Microfluidics based point-of-care diagnostics,” Biotechnol. J. 13(1), 1700047 (2018). https://doi.org/10.1002/biot.201700047). In addition, microfluidic techniques have found applications in patterning of cells and extracellular matrix (ECM) for applications in drug discovery and biomedicine (Nie et al., 202039. Nie, J., Fu, J., and He, Y., “Hydrogels: The next generation body materials for microfluidic chips?,” Small 16(46), 2003797 (2020). https://doi.org/10.1002/smll.202003797). There are multiple methods for fabrication of microfluidic devices; among those xurography is a low-cost and rapid fabrication technique, which does not require clean room facilities. Xurography, as a microfabrication method, was first introduced by Bartholomeusz et al. in 2005 (Bartholomeusz et al., 20055. Bartholomeusz, D. A., Boutté, R. W., and Andrade, J. D., “Xurography: Rapid prototyping of microstructures using a cutting plotter,” J. Microelectromech. Syst. 14(6), 1364–1374 (2005). https://doi.org/10.1109/JMEMS.2005.859087). In xurography, a physical blade is used to cut the materials based on a desirable design. Each layer of the device is individually cut. Then, the layers are aligned and laminated one after another to create a multilayer stack, which forms the microfluidic device (Mohammadzadeh et al., 201835. Mohammadzadeh, A., Fox-Robichaud, A. E., and Selvaganapathy, P. R., “Rapid and inexpensive method for fabrication of multi-material multi-layer microfluidic devices,” J. Micromech. Microeng. 29(1), 015013 (2018). https://doi.org/10.1088/1361-6439/aaf25a). Various materials including pressure sensitive adhesives (PSAs) (Islam et al., 201523. Islam, M., Natu, R., and Martinez-Duarte, R., “A study on the limits and advantages of using a desktop cutter plotter to fabricate microfluidic networks,” Microfluid. Nanofluid. 19(4), 973–985 (2015). https://doi.org/10.1007/s10404-015-1626-9; Mohammadzadeh et al., 201835. Mohammadzadeh, A., Fox-Robichaud, A. E., and Selvaganapathy, P. R., “Rapid and inexpensive method for fabrication of multi-material multi-layer microfluidic devices,” J. Micromech. Microeng. 29(1), 015013 (2018). https://doi.org/10.1088/1361-6439/aaf25a; and Yuen53. Yuen, P. K. and Goral, V. N., “Low-cost rapid prototyping of flexible microfluidic devices using a desktop digital craft cutter,” Lab Chip 10(3), 384–387 (2010). https://doi.org/10.1039/B918089C and Goral, 201053. Yuen, P. K. and Goral, V. N., “Low-cost rapid prototyping of flexible microfluidic devices using a desktop digital craft cutter,” Lab Chip 10(3), 384–387 (2010). https://doi.org/10.1039/B918089C), polymer films (Cassano et al., 20158. Cassano, C. L., Simon, A. J., Liu, W., Fredrickson, C., and Hugh Fan, Z., “Use of vacuum bagging for fabricating thermoplastic microfluidic devices,” Lab Chip 15(1), 62–66 (2015). https://doi.org/10.1039/C4LC00927D and Mohammadzadeh et al., 201835. Mohammadzadeh, A., Fox-Robichaud, A. E., and Selvaganapathy, P. R., “Rapid and inexpensive method for fabrication of multi-material multi-layer microfluidic devices,” J. Micromech. Microeng. 29(1), 015013 (2018). https://doi.org/10.1088/1361-6439/aaf25a), thin metal films (Mohammadzadeh et al., 201936. Mohammadzadeh, A., Robichaud, A. E. F., and Selvaganapathy, P. R., “Rapid and inexpensive method for fabrication and integration of electrodes in microfluidic devices,” J. Microelectromech. Syst. 28(4), 597–605 (2019). https://doi.org/10.1109/JMEMS.2019.2914110 and Stojanović et al., 201947. Stojanović, G., Paroški, M., Samardžić, N., Radovanović, M., and Krstić, D., “Microfluidics-based four fundamental electronic circuit elements resistor, inductor, capacitor and memristor,” Electronics 8(9), 960 (2019). https://doi.org/10.3390/electronics8090960), papers (Glavan et al., 201318. Glavan, A. C., Martinez, R. V., Maxwell, E. J., Subramaniam, A. B., Nunes, R. M., Soh, S., and Whitesides, G. M., “Rapid fabrication of pressure-driven open-channel microfluidic devices in omniphobic RF paper,” Lab Chip 13(15), 2922–2930 (2013). https://doi.org/10.1039/c3lc50371b and Samae et al., 202044. Samae, M., Ritmetee, P., Chirasatitsin, S., Kojić, S., Kojić, T., Jevremov, J., Stojanović, G., and Al Salami, H., “Precise manufacturing and performance validation of paper-based passive microfluidic micromixers,” Int. J. Precision Eng. Manuf. 21(3), 499–508 (2020). https://doi.org/10.1007/s12541-019-00272-0), and membranes (Fang et al., 201414. Fang, X., Wei, S., and Kong, J., “Based microfluidics with high resolution, cut on a glass fiber membrane for bioassays,” Lab Chip 14(5), 911–915 (2014). https://doi.org/10.1039/c3lc51246k) can be used as constituent layers in this fabrication method.Hydrogels are extensively used in the biomedical field as they are biocompatible, high permeable for small molecules, and optically clear. Also, some of the hydrogels can resemble biological tissues and extracellular matrix properties (Goy et al., 201919. Goy, C. B., Chaile, R. E., and Madrid, R. E., “Microfluidics and hydrogel: A powerful combination,” React. Funct. Polym. 145, 104314 (2019). https://doi.org/10.1016/j.reactfunctpolym.2019.104314; Nie et al., 202039. Nie, J., Fu, J., and He, Y., “Hydrogels: The next generation body materials for microfluidic chips?,” Small 16(46), 2003797 (2020). https://doi.org/10.1002/smll.202003797; and Zhang et al., 201654. Zhang, X., Li, L., and Luo, C., “Gel integration for microfluidic applications,” Lab Chip 16(10), 1757–1776 (2016). https://doi.org/10.1039/C6LC00247A). Hydrogels have been used as barriers for increasing fluid resistance and creating a free diffusion environment in microfluidic devices (Zhang et al., 201654. Zhang, X., Li, L., and Luo, C., “Gel integration for microfluidic applications,” Lab Chip 16(10), 1757–1776 (2016). https://doi.org/10.1039/C6LC00247A). They have been incorporated into microfluidic devices used for various applications including cell culture, biosensors, gradient generator, as well as in creation of active elements such as valves and separation devices. Hydrogels are a vital part of cell culture and cell patterning microdevices (Gao et al., 201015. Gao, D., Liu, J., Wei, H.-B., Li, H.-F., Guo, G.-S., and Lin, J.-M., “A microfluidic approach for anticancer drug analysis based on hydrogel encapsulated tumor cells,” Anal. Chim. Acta 665(1), 7–14 (2010). https://doi.org/10.1016/j.aca.2010.03.015 and Koh et al., 200326. Koh, W.-G., Itle, L. J., and Pishko, M. V., “Molding of hydrogel microstructures to create multiphenotype cell microarrays,” Anal. Chem. 75(21), 5783–5789 (2003). https://doi.org/10.1021/ac034773s). Furthermore, they are able to keep biomolecules inside and form a 3D environment for optical (Duan et al., 201711. Duan, K., Ghosh, G., and Lo, J. F., “Optimizing multiplexed detections of diabetes antibodies via quantitative microfluidic droplet array,” Small 13(46), 1702323 (2017). https://doi.org/10.1002/smll.201702323 and Jung et al., 201524. Jung, Y. K., Kim, J., and Mathies, R. A., “Microfluidic linear hydrogel array for multiplexed single nucleotide polymorphism (SNP) detection,” Anal. Chem. 87(6), 3165–3170 (2015). https://doi.org/10.1021/ac5048696) or electrochemical (Aymerich et al., 20183. Aymerich, J., Márquez, A., Terés, L., Muñoz-Berbel, X., Jiménez, C., Domínguez, C., Serra-Graells, F., and Dei, M., “Cost-effective smartphone-based reconfigurable electrochemical instrument for alcohol determination in whole blood samples,” Biosens. Bioelectron. 117, 736–742 (2018). https://doi.org/10.1016/j.bios.2018.06.044 and Matharu et al., 201332. Matharu, Z., Enomoto, J., and Revzin, A., “Miniature enzyme-based electrodes for detection of hydrogen peroxide release from alcohol-injured hepatocytes,” Anal. Chem. 85(2), 932–939 (2013). https://doi.org/10.1021/ac3025619) biosensors. Hydrogel permeability to diffusion has been exploited in gradient generator microdevices (Bachmann et al., 20184. Bachmann, B., Spitz, S., Rothbauer, M., Jordan, C., Purtscher, M., Zirath, H., Schuller, P., Eilenberger, C., Ali, S. F., and Mühleder, S., “Engineering of three-dimensional pre-vascular networks within fibrin hydrogel constructs by microfluidic control over reciprocal cell signaling,” Biomicrofluidics 12(4), 042216 (2018). https://doi.org/10.1063/1.5027054; Wu et al., 201550. Wu, X., Newbold, M. A., and Haynes, C. L., “Recapitulation of in vivo-like neutrophil transendothelial migration using a microfluidic platform,” Analyst 140(15), 5055–5064 (2015). https://doi.org/10.1039/C5AN00967G; and Yoon et al., 201652. Yoon, D., Kim, H., Lee, E., Park, M. H., Chung, S., Jeon, H., Ahn, C.-H., and Lee, K., “Study on chemotaxis and chemokinesis of bone marrow-derived mesenchymal stem cells in hydrogel-based 3D microfluidic devices,” Biomater. Res. 20(1), 1–8 (2016). https://doi.org/10.1186/s40824-016-0070-6). Some classes of hydrogels can undergo volume transition in response to external stimuli such as temperature, pH, and light (Ahmed, 20151. Ahmed, E. M., “Hydrogel: Preparation, characterization, and applications: A review,” J. Adv. Res. 6(2), 105–121 (2015). https://doi.org/10.1016/j.jare.2013.07.006). These types of hydrogels have been used to create active elements such as valves and actuators inside a microchannel (Beebe et al., 20007. Beebe, D. J., Moore, J. S., Bauer, J. M., Yu, Q., Liu, R. H., Devadoss, C., and Jo, B.-H., “Functional hydrogel structures for autonomous flow control inside microfluidic channels,” Nature 404(6778), 588–590 (2000). https://doi.org/10.1038/35007047 and Harmon et al., 200320. Harmon, M. E., Tang, M., and Frank, C. W., “A microfluidic actuator based on thermoresponsive hydrogels,” Polymer 44(16), 4547–4556 (2003). https://doi.org/10.1016/S0032-3861(03)00463-4). Most importantly, hydrogels have been an essential part of many separation microfluidic devices, such as cell separation (Lin et al., 201528. Lin, R.-Z., Hatch, A., Antontsev, V. G., Murthy, S. K., and Melero-Martin, J. M., “Microfluidic capture of endothelial colony-forming cells from human adult peripheral blood: Phenotypic and functional validation in vivo,” Tissue Eng. Part C: Methods 21(3), 274–283 (2015). https://doi.org/10.1089/ten.tec.2014.0323), electrophoretic concentration and separation of biomolecules (Gerver17. Gerver, R. E. and Herr, A. E., “Microfluidic western blotting of low-molecular-mass proteins,” Anal. Chem. 86(21), 10625–10632 (2014). https://doi.org/10.1021/ac5024588 and Herr, 201417. Gerver, R. E. and Herr, A. E., “Microfluidic western blotting of low-molecular-mass proteins,” Anal. Chem. 86(21), 10625–10632 (2014). https://doi.org/10.1021/ac5024588 and Mohamadi et al., 201534. Mohamadi, R. M., Svobodova, Z., Bilkova, Z., Otto, M., Taverna, M., Descroix, S., and Viovy, J.-L., “An integrated microfluidic chip for immunocapture, preconcentration and separation of β-amyloid peptides,” Biomicrofluidics 9(5), 054117 (2015). https://doi.org/10.1063/1.4931394), and isoelectric focusing separation microdevices (Herzog et al., 201622. Herzog, C., Poehler, E., Peretzki, A. J., Borisov, S. M., Aigner, D., Mayr, T., and Nagl, S., “Continuous on-chip fluorescence labelling, free-flow isoelectric focusing and marker-free isoelectric point determination of proteins and peptides,” Lab Chip 16(9), 1565–1572 (2016). https://doi.org/10.1039/C6LC00055J and Poehler et al., 201542. Poehler, E., Herzog, C., Lotter, C., Pfeiffer, S. A., Aigner, D., Mayr, T., and Nagl, S., “Label-free microfluidic free-flow isoelectric focusing, pH gradient sensing and near real-time isoelectric point determination of biomolecules and blood plasma fractions,” Analyst 140(22), 7496–7502 (2015). https://doi.org/10.1039/C5AN01345C).Multiple methods have been used for integration hydrogels into microfluidic devices such as gel photopolymerization for creating micro-structures, and flow-solidification based integration (Zhang et al., 201654. Zhang, X., Li, L., and Luo, C., “Gel integration for microfluidic applications,” Lab Chip 16(10), 1757–1776 (2016). https://doi.org/10.1039/C6LC00247A). In these methods, hydrogel is mostly integrated in-plane and in the same level of the microfluidic channels, not as a separate layer. In flow-solidification based method (Chung et al., 200910. Chung, S., Sudo, R., Mack, P. J., Wan, C.-R., Vickerman, V., and Kamm, R. D., “Cell migration into scaffolds under co-culture conditions in a microfluidic platform,” Lab Chip 9(2), 269–275 (2009). https://doi.org/10.1039/B807585A; Luo et al., 201031. Luo, X., Shen, K., Luo, C., Ji, H., Ouyang, Q., and Chen, Y., “An automatic microturbidostat for bacterial culture at constant density,” Biomed. Microdevices 12(3), 499–503 (2010). https://doi.org/10.1007/s10544-010-9406-5; and Puchberger-Enengl et al., 201443. Puchberger-Enengl, D., Krutzler, C., Keplinger, F., and Vellekoop, M. J., “Single-step design of hydrogel-based microfluidic assays for rapid diagnostics,” Lab Chip 14(2), 378–383 (2014). https://doi.org/10.1039/C3LC50944C), the gel solidification is controlled by surface tension. For instance, hydrophilic patterns on a hydrophobic surface were used to control the localize where the gel was introduced (Zhang et al., 201654. Zhang, X., Li, L., and Luo, C., “Gel integration for microfluidic applications,” Lab Chip 16(10), 1757–1776 (2016). https://doi.org/10.1039/C6LC00247A). However, this method is not always straightforward and can add complexity to fabrication.Photopolymerization of the UV-curable gels can be used for embedding hydrogels in the microfluidic devices as well (Beck et al., 20206. Beck, A., Obst, F., Busek, M., Grünzner, S., Mehner, P. J., Paschew, G., Appelhans, D., Voit, B., and Richter, A., “Hydrogel patterns in microfluidic devices by do-it-yourself UV-photolithography suitable for very large-scale integration,” Micromachines 11(5), 479 (2020). https://doi.org/10.3390/mi11050479; Beebe et al., 20007. Beebe, D. J., Moore, J. S., Bauer, J. M., Yu, Q., Liu, R. H., Devadoss, C., and Jo, B.-H., “Functional hydrogel structures for autonomous flow control inside microfluidic channels,” Nature 404(6778), 588–590 (2000). https://doi.org/10.1038/35007047; Chen et al., 20179. Chen, L., Wang, K. X., and Doyle, P. S., “Effect of internal architecture on microgel deformation in microfluidic constrictions,” Soft Matter 13(9), 1920–1928 (2017). https://doi.org/10.1039/C6SM02674E; Garcia-Schwarz16. Garcia-Schwarz, G. and Santiago, J. G., “Integration of on-chip isotachophoresis and functionalized hydrogels for enhanced-sensitivity nucleic acid detection,” Anal. Chem. 84(15), 6366–6369 (2012). https://doi.org/10.1021/ac301586q and Santiago, 201216. Garcia-Schwarz, G. and Santiago, J. G., “Integration of on-chip isotachophoresis and functionalized hydrogels for enhanced-sensitivity nucleic acid detection,” Anal. Chem. 84(15), 6366–6369 (2012). https://doi.org/10.1021/ac301586q; Jung et al., 201625. Jung, Y. K., Kim, J., and Mathies, R. A., “Microfluidic hydrogel arrays for direct genotyping of clinical samples,” Biosens. Bioelectron. 79, 371–378 (2016). https://doi.org/10.1016/j.bios.2015.12.068; Koh27. Koh, W.-G. and Pishko, M., “Immobilization of multi-enzyme microreactors inside microfluidic devices,” Sens. Actuators, B 106(1), 335–342 (2005). https://doi.org/10.1016/j.snb.2004.08.025 and Pishko, 200527. Koh, W.-G. and Pishko, M., “Immobilization of multi-enzyme microreactors inside microfluidic devices,” Sens. Actuators, B 106(1), 335–342 (2005). https://doi.org/10.1016/j.snb.2004.08.025; Liu et al., 200929. Liu, J., Gao, D., Li, H.-F., and Lin, J.-M., “Controlled photopolymerization of hydrogel microstructures inside microchannels for bioassays,” Lab Chip 9(9), 1301–1305 (2009). https://doi.org/10.1039/b819219g,; Nash37. Nash, A. T., Foster, D. A., Thompson, S. I., Han, S., Fernandez, M. K., and Hwang, D. K., “A new rapid microfluidic detection platform utilizing hydrogel-membrane under cross-flow,” Adv. Mater. Technol. 7, 2101396 (2022). https://doi.org/10.1002/admt.202101396 and Foster, 202237. Nash, A. T., Foster, D. A., Thompson, S. I., Han, S., Fernandez, M. K., and Hwang, D. K., “A new rapid microfluidic detection platform utilizing hydrogel-membrane under cross-flow,” Adv. Mater. Technol. 7, 2101396 (2022). https://doi.org/10.1002/admt.202101396; Nguyen et al., 202038. Nguyen, H.-T., Massino, M., Keita, C., and Salmon, J.-B., “Microfluidic dialysis using photo-patterned hydrogel membranes in PDMS chips,” Lab Chip 20(13), 2383–2393 (2020). https://doi.org/10.1039/D0LC00279H; Paustian et al., 201341. Paustian, J. S., Azevedo, R. N., Lundin, S.-T. B., Gilkey, M. J., and Squires, T. M., “Microfluidic microdialysis: Spatiotemporal control over solution microenvironments using integrated hydrogel membrane microwindows,” Phys. Rev. X 3(4), 041010 (2013). https://doi.org/10.1103/PhysRevX.3.041010). Uniform exposure, using a mask, or direct writing are the methods used for photopolymerization. Some UV-curable hydrogels that have been incorporated into microfluidic devices include polyethylene glycol diacrylate (PEG-DA), poly-N-isopropyl acrylamide (pNIPAAM), and polyacrylamide (PAAM) (Zhang et al., 201654. Zhang, X., Li, L., and Luo, C., “Gel integration for microfluidic applications,” Lab Chip 16(10), 1757–1776 (2016). https://doi.org/10.1039/C6LC00247A). This technique provides desired features with high resolution; however, UV-curable hydrogels are limited, and they are difficult to integrate in post-processing as they require the unpolymerized gel to be removed from the channels. In a typical fabrication process using this technique, the microchannels are first filled with a precursor. In the next step, the desired regions are photopolymerized and the uncrosslinked gel precursor is removed. These post-polymerization steps make this technique relatively complicated (Eker et al., 201413. Eker, B., Temiz, Y., and Delamarche, E., “Heterogeneous integration of gels into microfluidics using a mesh carrier,” Biomed. Microdevices 16(6), 829–835 (2014). https://doi.org/10.1007/s10544-014-9886-9 and Heo and Crooks, 200521. Heo, J. and Crooks, R. M., “Microfluidic biosensor based on an array of hydrogel-entrapped enzymes,” Anal. Chem. 77(21), 6843–6851 (2005). https://doi.org/10.1021/ac0507993). Eker et al. used mesh carriers to transfer and embed small volumes of a biomolecule loaded poly(ethylene)glycol-based acrylamide (PEGACA) hydrogel in the desired locations of microfluidic devices to serve as reagent reservoirs. The mesh was used as a transfer substrate instead of a scaffold and was not integrated into the device. Furthermore, the gel was photopolymerized in the form of drops and transferred to the microfluidic chip without any functional assessment such as its ability to seal or retain shape upon drying and reconstitution (Eker et al., 201413. Eker, B., Temiz, Y., and Delamarche, E., “Heterogeneous integration of gels into microfluidics using a mesh carrier,” Biomed. Microdevices 16(6), 829–835 (2014). https://doi.org/10.1007/s10544-014-9886-9). In summary, most of the methods in the literature require the integration of gels as a post-processing step that will add extra cost to the fabrication of the device. They are also more suited to in-plane fabrication of gels in microfluidic channels such as in DNA or protein separation devices. Very few methods exist that can integrate gels as part of the fabrication process flow. None of these methods are suited in applications where the gel needs to be integrated in-between two channel layers either as a diffusion barrier in diagnostic applications or to hold cells exposed to different analytes on either side in organ on chip type applications. Finally, drying and rehydration of the gels and the consequent effect on their integrity and leakage have not been studied in the literature but is crucial for devices that are to be stored for up to 6–12 month prior to use in the field.

Here, a low-cost and easy to implement method to integrate hydrogels into microfluidic devices during the fabrication process is introduced to overcome these limitations. It is shown that thin porous and fibrous membranes, such as electrospun PCL membrane can be used as a scaffold for hydrogel embedding within microfluidic devices. In this method, the hydrogel is integrated as a separate layer, sandwiched between other layers of the device. Xurography is used to expose regions of the membrane onto which the hydrogels can be loaded. The same technique has been used to integrate the gel loaded membranes into microfluidic channels. It is also shown that PCL membrane can prevent complete collapse of the gel upon dehydration and retain the macrostructure of hydrogel after rehydration. It also maintains intimate contact between the gel and the sidewalls of the channels which is important in preventing leakages and convective flows. These features make this hybrid approach suitable for long term dried storage of hydrogels inside microfluidic devices possible for the first time, thereby avoiding loading of gels at the point of use. A demonstration of its usefulness was performed by fabricating a point of care microfluidic device containing a dried agarose loaded membrane for accumulation and quantification of 150 bp DNA, which is useful for developing a tool for sepsis prognostics.

II. EXPERIMENTAL

Section:

ChooseTop of pageABSTRACTI. INTRODUCTIONII. EXPERIMENTAL <<III. RESULTS AND DISCUSSI...IV. CONCLUSIONSUPPLEMENTARY MATERIALREFERENCES

A. Materials

Transfer adhesive tape (7952MP-3M™, 50 μm thickness) was obtained from 3M™, Maplewood, MN. A one-sided hydrophilic adhesive tape (polyester film with acrylic adhesive (93049-AR), 100 μm thickness) was obtained from adhesive research (AR), Glen Rock, PA. Polyvinyl chloride (PVC) films (Clear-Lay, 127 μm thickness) were from Grafix, Maple Heights, OH. A polyimide (Kapton) tape with a silicone adhesive (double-sided and double Liner, 100 μm) was from Caplinq. The copper-polyimide composite foil (9 μm-Cu and 12 μm-PI thickness, Pyralux®) was acquired from DuPont, USA. The dicing tape (UV curable acrylic adhesive on PVC film, 95 μm thickness) was obtained from Ultron Systems, US. A Whatman cellulose chromatography paper (1 Chr, 180 μm thickness) was from Sigma Aldrich. Agarose powder and 50× TAE buffer concentrate were from BioShop Canada Inc, Burlington, ON. Polycaprolactone average Mn 80 000, dichloromethane and methylene blue were from Sigma Aldrich. Quant-iT PicoGreen® dsDNA kit was from Life Technologies. A 150 bp DNA sample was provided by Thrombosis & Atherosclerosis Research Institute (TaARI).

B. Fabrication process

1. Electrospinning and membrane fabrication

A non-woven porous polycaprolactone (PCL) membrane was prepared by electrospinning. Briefly, a 10 wt./vol.% PCL solution in a mixture of dichloromethane (DCM) and ethanol with a 4:1 volume ratio was prepared. The electrospinning was performed at optimized parameters of 13 kV applied voltage, 8 cm distance between the needle (blunt needle gauge 18) and the aluminum collector (a flat stationary surface), and flow rate of 0.3 ml/h to produce uniform and narrow distribution of fiber diameter and bead free fibers. The process was performed under ambient conditions at room temperature. Electrospinning was performed for 25 min which resulted in a 250 ± 39.8 μm thick membrane [number of measurements (n) = 20]. SEM image of the electrospun PCL membrane is presented in Fig. 1(a). Fiber size was measured from the SEM images of over 100 fibers using ImageJ. Figure 1(b) demonstrates the fiber diameter distribution. Fibers diameter average size was 1.05 ± 0.27 μm.

2. Hydrogel embedding into the membrane

The electrospun PCL membrane is hydrophobic and has a contact angle of 112 ± 3 (n = 3), measured using an optical goniometer (OCA 35). Therefore, embedding hydrogels into it is a challenge. The hydrophilicity of the regions of PCL membrane intended for the integration of the hydrogel was increased selectively by plasma treatment using the process shown in Fig. 2. Two layers of double-sided Kapton tape were cut using a cutter plotter (FC8600, Graphtec America, US) based on the desired designs for embedded hydrogels in the device [Fig. 2(a)]. Layers were assembled in a layer-by-layer manner with a proper alignment on either side of the PCL layer in the center. Next, this sandwiched layer was exposed to oxygen plasma for 80 s [Fig. 2(b)] which selectively makes the exposed regions of the PCL membrane hydrophilic (contact angle ∼0°) while the rest of the membrane remains hydrophobic. Then, the hydrogel (2 wt.% agarose in the TAE buffer, prepared by microwaving the solution) was pipeted in the open hydrophilic regions on the membrane and its excess amount was removed from the surface after gelation [Fig. 2(c)].

3. Drying and rehydration of embedded hydrogels

The agarose loaded electrospun PCL membrane was dried overnight at room temperature. The next day, one droplet of 1× TAE buffer was used for rehydration of agarose in the membrane for 1 minute. A camera (Infinity3, Lumenera, US) mounted on a stereo microscope (SZ61, Olympus, Japan) was used to take images of cross section of the membrane before and after rehydration. SEM microscopy was also used to further evaluate the microstructure of the dried agarose embedded membrane.

4. Evaluation of the sealing of the hydrogel by the transport of colorimetric dye

A device containing two cross channels was fabricated using the xurography method. The schematic of the layers of the device and the exploded view of the layers are shown in Figs. 3(a) and 3(b), respectively. A cutter plotter was used to cut different layers of the final device in predefined patterns. Circular openings with 1.1 mm diameter for the hydrogel section were cut into two layers of the Kapton tape. Assembly of the layers and hydrogel integration process was similar as before (in the case of two and three layers of PCL, more membrane layers were stacked on each other between two Kapton tapes and plasma exposure and hydrogel loading were performed from both sides of membrane layer). Two channels with 1.6 mm width and 10 mm length were cut out of 127 μm thick PVC films and were attached to the top and bottom of the Kapton tapes. Finally, a hydrophilic adhesive (93049-AR) with openings for inlets and outlets was attached to the top channel. Another layer of hydrophilic adhesive was used to seal the bottom channel. A cross-sectional view of the device with dimensions with respect to the scale of the different layers is demonstrated in Fig. 3(c). All the layers were aligned one after the other using alignment marks. At the end, the device passed through a laminator.

The fabricated device with intersecting channels was used to study the sealing of the hydrated and rehydrated agarose in the membrane. Top and bottom channels were filled with 1× TAE buffer and methylene blue solution, respectively. The diffusion of methylene blue from bottom to top channel was tracked using a camera (Infinity3, Lumenera, US) mounted on a stereo microscope (SZ61, Olympus, Japan). Images were taken every 4 min and were analyzed using ImageJ. The original RGB images were transferred to a grey scale format ranging from 0 to 255 and the intensity in the circular agarose region was calculated as an aggregate grey scale value. The brighter color gives a higher grey scale value. The intensity was normalized by subtracting the obtained grey scale value from the starting grey value (background) for each image before the dye was added.

The effect of electric field on the transport of methylene blue was studied by incorporating electrodes in the fabricated device using xurography. Electrodes were made from a thin copper film (21 μm thickness) and dicing tape was used as the support for electrode patterns. Copper foil was sandwiched between two layers of dicing tape. Then, the top layer and metal film were cut using the cutter plotter (top dicing tape prevents the metal from being torn in the process of cutting). Then, the top and bottom sides were exposed to UV light to make the dicing tape non-sticky and help with peeling the unwanted regions of metal foil. On top of the electrode layer, a transfer adhesive tape was used to attach the electrodes to the other layers of the device [Fig. 4(a)]. The channel containing methylene blue and the buffer channel were connected to anode and cathode, respectively. A 10 V was applied using a DC power supply (Keithley 2636) for 6 min between the two microchannels and images of the hydrogel region were taken to analyze the mass transfer. The sealing efficiency of the dried (air-dried overnight) and rehydrated agarose in the membrane was studied similarly. First, the hydrogel was rehydrated using TAE buffer in the top channel for 1 min. Then, the bottom channel was filled with methylene blue and transport of that in the absence and presence of electric field was observed.

5. Concentration and quantification of DNA in agarose loaded membranes

A cross channel microfluidic device [Fig. 4(b)] was fabricated to concentrate and quantify DNA from its initial solution. The devices used for the DNA concentration experiments were fabricated similar to those in the previous section. The hydrogel region was defined using the PCL membrane which had been sandwiched between two Kapton tapes with circular openings of 350 μm radius in one layer and 700 μm in the other. The sample channel was fabricated with dimensions of 800 μm (W), 127 μm (H), and 8 mm (L) and buffer channel with dimensions of 1.6 mm (W), 127 μm (H), and 8 mm (L). The open region was filled with 2% agarose after exposure to oxygen plasma. The agarose embedded layer dried overnight at room temperature. Device was sealed with a hydrophilic tape (93049-AR) containing the inlets and outlets of the channels.DNA samples were prepared by diluting the 150 bp DNA stock solution in 1× TE (200 mM Tris-HCl and 20 mM EDTA) buffer to the desired concentrations (1 and 5 μg/ml). Then, 200 times diluted PicoGreen was added to the DNA sample with a 1:1 volume ratio. The tube containing prepared sample was kept in dark for 10 min for the DNA and dye to completely intercalated. A fluorescent microscope (Etaluma, Model 500) was used for imaging purposes. After connecting the sample channel and buffer channel inlets to cathode and anode, respectively, the buffer channel was filled with 4 μl of 1× TAE buffer to rehydrate agarose embedded in the membrane. One minute later, the sample channel was filled with 2 μl DNA sample. Cross section of the channels is presented in Fig. 4(c). The electrodes were connected to a DC power supply (Keithley 2636) and a 5 V voltage was applied to the device to initiate the DNA transfer.

III. RESULTS AND DISCUSSION

Section:

ChooseTop of pageABSTRACTI. INTRODUCTIONII. EXPERIMENTALIII. RESULTS AND DISCUSSI... <<IV. CONCLUSIONSUPPLEMENTARY MATERIALREFERENCES

A. Hydrogel integration, resolution characterization, and patterning

Electrospinning has been used here to create a non-woven fibrous membrane with large pores and high porosity to serve as a scaffold to embed a small volume of hydrogel. The use of the scaffold is beneficial in maintaining the macrostructure of the hydrogel even when it is dried and reconstituted, which is important for many point-of-care applications.

The electrospun membrane was made from 10 wt./vol.% PCL solution in a mixture of DCM and ethanol. Various applied voltage and distance between the needle and collector were used to arrive at the final optimized conditions for electrospinning. An applied voltage was 13 kV and the distance between the needle and the collector of 8 cm with a solution flow rate of 0.3 ml/h was found to be optimal to produce uniform fiber diameter, pore size, and bead free membranes. Electrospinning was performed for 25 min in order to achieve a 250 ± 39.8 μm thick membrane with average 1.05 ± 0.27 μm fiber diameter size. The fabricated membrane was suitable for embedding enough amount of hydrogel, while the structure of hydrogel is maintained by fibers. Thinner membranes proved to be more fragile and less mechanically stable when loaded with the hydrogel.

The as spun PCL membrane is hydrophobic (contact angle of 112 ± 3) which is suitable to prevent spread of the imbibed gel further from the patterned regions. However, it also prevents complete and robust imbibition of the gel within the patterned regions. Therefore, a self-aligned hydrophilization process was developed that can convert the patterned regions of the membrane for robust imbibition of the gel. It has been shown that surface plasma treatment enhances the hydrophilicity of the PCL membrane by forming oxygen-containing groups (Mochane et al., 201933. Mochane, M. J., Motsoeneng, T. S., Sadiku, E. R., Mokhena, T. C., and Sefadi, J. S., “Morphology and properties of electrospun PCL and its composites for medical applications: A mini review,” Appl. Sci. 9(11), 2205 (2019). https://doi.org/10.3390/app9112205). Therefore, PCL membrane was sandwiched between two Kapton tape layers with opening sections for hydrogel loading. These Kapton tape layers protect the PCL membrane from exposure to plasma and consequent hydrophilization in all the regions except in the openings where the hydrogel is to be loaded. Upon dropping a small bolus of hydrogel, the plasma treated region also imbibes the gel while the surrounding regions do not due to their hydrophobic nature. The excess gel on top of the membrane was wiped away leaving behind a precisely defined volume of hydrogel that was loaded on to the membrane. For demonstrations, we use agarose as the hydrogel, but this method should be applicable to any other hydrogel as well. Hydrophilic regions of the membrane were filled with agarose colored with food dyes.Using this technique, hydrogel microstructures in different shapes and sizes can be obtained. Thickness of the hydrogel layer is controlled by the thickness of the membrane. By making the membrane thicker or stacking several membranes on top of each other, a thicker gel layer can be obtained. In order to determine the lateral resolution of this method, a series of circular and square regions with different sizes were cut and filled with hydrogel. Circular and square patterns (ranging from 100 μm to 1 mm with 100 μm increments) were fabricated and filled with 2% agarose colored with food dyes. It is shown that different patterns of hydrogel with dimensions as small as 300 μm can be created (Fig. 5). This resolution corresponds to ∼ 20 nl of the hydrogel inside the membrane (considering thickness of the membrane be 250 μm and ignoring the volume of the fibers). Xurography was not able to create reliable patterns smaller than 300 μm in the Kapton tape and the patterns did not resemble the designed shapes below that limit. Although this resolution is less than the resolution associated with the photopolymerization method for creating gel microstructures (25 μm) (Beebe et al., 20007. Beebe, D. J., Moore, J. S., Bauer, J. M., Yu, Q., Liu, R. H., Devadoss, C., and Jo, B.-H., “Functional hydrogel structures for autonomous flow control inside microfluidic channels,” Nature 404(6778), 588–590 (2000). https://doi.org/10.1038/35007047), it is still sufficient for many point of care applications where hydrogel integration is desired. This hydrogel integration technique resolution can be improved by using a better cutter plotter machine with higher resolutions. Moreover, using a thinner double-sided tape (Kapton tape thickness is 250 μm) should be helpful for getting better cuts and greater resolution. Alternatively, laser micromachining can be used to cut the Kapton tape with a much higher resolution.The xurographic process is flexible and can create a wide range of patterns. Long line type patterns are important for creating separation columns from gels and can also be patterned using the xurographic gel integration process. As shown in Fig. 6(a), long hydrogel lines with width as small as 200 μm can be created. Xurography did not provide a good cut for 150 μm wide line. It is also demonstrated that by using this technique more complex patterns such as alphabets shown in Fig. 6(b) can be obtained. Another benefit of this technique is its scalability. The xurographic process can be used to create multiple gel locations on the same device. These locations can be imbibed with different gel compositions that can be used to create a microarray as shown in Fig. 6(c). This capability is especially advantageous when multiple conditions, concentrations, and analytes need to be detected or analyzed using the same sample. The layer containing hydrogel patterns can be attached by adhesives to the other layers of microfluidic devices fabricated using the xurography technique. This method allows for the integration of gels into microfluidic devices in an out-of-plane manner. It also allows the integration of gels in-between two microfluidic channels where it could be used as a con